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Removal of visual cortex in the rat axotomizes projection neurons in the dorsal lateral geniculate nucleus (dLGN), leading to cytological and structural changes and apoptosis. Biotinylated dextran amine was injected into the visual cortex to label dLGN projection neurons retrogradely prior to removing the cortex in order to quantify the changes in the dendritic morphology of these neurons that precede cell death. At 12 hours after axotomy we observed a loss of appendages and the formation of varicosities in the dendrites of projection neurons. During the next 7 days, the total number of dendrites and the cross-sectional areas of the dendritic arbors of projection neurons declined to about 40% and 20% of normal, respectively. The response of dLGN projection neurons to axotomy was asynchronous, but the sequence of structural changes in individual neurons was similar; namely, disruption of dendrites began within hours followed by cell soma atrophy and nuclear condensation that commenced after the loss of secondary dendrites had occurred. However, a single administration of fibroblast growth factor-2 (FGF2), which mitigates injury-induced neuronal cell death in the dLGN when given at the time of axotomy, markedly reduced the dendritic degeneration of projection neurons. At 3 and 7 days after axotomy the number of surviving dendrites of dLGN projection neurons in FGF-2 treated rats was approximately 50% greater than in untreated rats, and the cross-sectional areas of dendritic arbors were approximately 60% and 50% larger. Caspase-3 activity in axotomized dLGN projection neurons was determined by immunostaining for fractin (fractin-IR), an actin cleavage product produced exclusively by activated caspase-3. Fractin-IR was seen in some dLGN projection neurons at 36 hours survival, and it increased slightly by 3 days. A marked increase in reactivity was seen by 7 days, with the entire dLGN filled with dense fractin-IR in neuronal cell somas and dendrites.

The rodent visual system encompasses retinal ganglion cells and their axons that form the optic nerve to enter thalamic and midbrain centers, and postsynaptic projections to the visual cortex. Based on its distinct anatomical structure and convenient accessibility, it has become the favored structure for studies on neuronal survival, axonal regeneration, and synaptic plasticity. Recent advancements in MR imaging have enabled the in vivo visualization of the retino-tectal part of this projection using manganese mediated contrast enhancement (MEMRI). Here, we present a MEMRI protocol for illustration of the visual projection in mice, by which resolutions of (200 µm)3 can be achieved using common 3 Tesla scanners. We demonstrate how intravitreal injection of a single dosage of 15 nmol MnCl2 leads to a saturated enhancement of the intact projection within 24 hr. With exception of the retina, changes in signal intensity are independent of coincided visual stimulation or physiological aging. We further apply this technique to longitudinally monitor axonal degeneration in response to acute optic nerve injury, a paradigm by which Mn2+ transport completely arrests at the lesion site. Conversely, active Mn2+ transport is quantitatively proportionate to the viability, number, and electrical activity of axon fibers. For such an analysis, we exemplify Mn2+ transport kinetics along the visual path in a transgenic mouse model (NF-κB p50KO) displaying spontaneous atrophy of sensory, including visual, projections. In these mice, MEMRI indicates reduced but not delayed Mn2+ transport as compared to wild type mice, thus revealing signs of structural and/or functional impairments by NF-κB mutations.
In summary, MEMRI conveniently bridges in vivo assays and post mortem histology for the characterization of nerve fiber integrity and activity. It is highly useful for longitudinal studies on axonal degeneration and regeneration, and investigations of mutant mice for genuine or inducible phenotypes.

Purkinje cells are an attractive model system for studying dendritic development, because they have an impressive dendritic tree which is strictly oriented in the sagittal plane and develops mostly in the postnatal period in small rodents 3. Furthermore, several antibodies are available which selectively and intensively label Purkinje cells including all processes, with anti-Calbindin D28K being the most widely used. For viewing of dendrites in living cells, mice expressing EGFP selectively in Purkinje cells 11 are available through Jackson labs. Organotypic cerebellar slice cultures cells allow easy experimental manipulation of Purkinje cell dendritic development because most of the dendritic expansion of the Purkinje cell dendritic tree is actually taking place during the culture period 4. We present here a short, reliable and easy protocol for viewing and analyzing the dendritic morphology of Purkinje cells grown in organotypic cerebellar slice cultures. For many purposes, a quantitative evaluation of the Purkinje cell dendritic tree is desirable. We focus here on two parameters, dendritic tree size and branch point numbers, which can be rapidly and easily determined from anti-calbindin stained cerebellar slice cultures. These two parameters yield a reliable and sensitive measure of changes of the Purkinje cell dendritic tree. Using the example of treatments with the protein kinase C (PKC) activator PMA and the metabotropic glutamate receptor 1 (mGluR1) we demonstrate how differences in the dendritic development are visualized and quantitatively assessed. The combination of the presence of an extensive dendritic tree, selective and intense immunostaining methods, organotypic slice cultures which cover the period of dendritic growth and a mouse model with Purkinje cell specific EGFP expression make Purkinje cells a powerful model system for revealing the mechanisms of dendritic development.

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve.
The optic nerve can be accessed within the orbit of the eye and completely transected (axotomized), cutting the axons of the entire RGC population. Optic nerve
transection is a reproducible model of apoptotic neuronal cell death in the adult CNS 1-4. This model is particularly attractive because the vitreous
chamber of the eye acts as a capsule for drug delivery to the retina, permitting experimental manipulations via intraocular injections. The diffusion of chemicals
through the vitreous fluid ensures that they act upon the entire RGC population. Moreover, RGCs can be selectively transfected by applying short interfering RNAs
(siRNAs), plasmids, or viral vectors to the cut end of the optic nerve 5-7 or injecting vectors into their target, the superior colliculus 8.
This allows researchers to study apoptotic mechanisms in the desired neuronal population without confounding effects on other bystander neurons or surrounding glia.
An additional benefit is the ease and accuracy with which cell survival can be quantified after injury. The retina is a flat, layered tissue and RGCs are localized in
the innermost layer, the ganglion cell layer. The survival of RGCs can be tracked over time by applying a fluorescent tracer (3% Fluorogold) to the cut end of the
optic nerve at the time of axotomy, or by injecting the tracer into the superior colliculus (RGC target) one week prior to axotomy. The tracer is retrogradely transported, labeling
the entire RGC population. Because the ganglion cell layer is a monolayer (one cell thick), RGC densities can be quantified in flat-mounted tissue, without the need
for stereology. Optic nerve transection leads to the apoptotic death of 90% of injured RGCs within 14 days postaxotomy 9-11. RGC apoptosis has a
characteristic time-course whereby cell death is delayed 3-4 days postaxotomy, after which the cells rapidly degenerate. This provides a time window for
experimental manipulations directed against pathways involved in apoptosis.

To understand how differences in complex cell shapes are achieved, it is important to accurately follow microtubule organization. The Drosophila larval body wall contains several cell types that are models to study cell and tissue morphogenesis. For example tracheae are used to examine tube morphogenesis1, and the dendritic arborization (DA) sensory neurons of the Drosophila larva have become a primary system for the elucidation of general and neuron-class-specific mechanisms of dendritic differentiation2-5 and degeneration6.
The shape of dendrite branches can vary significantly between neuron classes, and even among different branches of a single neuron7,8. Genetic studies in DA neurons suggest that differential cytoskeletal organization can underlie morphological differences in dendritic branch shape4,9-11. We provide a robust immunological labeling method to assay in vivo microtubule organization in DA sensory neuron dendrite arbor (Figures 1, 2, Movie 1). This protocol illustrates the dissection and immunostaining of first instar larva, a stage when active sensory neuron dendrite outgrowth and branching organization is occurring 12,13.
In addition to staining sensory neurons, this method achieves robust labeling of microtubule organization in muscles (Movies 2, 3), trachea (Figure 3, Movie 3), and other body wall tissues. It is valuable for investigators wishing to analyze microtubule organization in situ in the body wall when investigating mechanisms that control tissue and cell shape.

Chronic Imaging of Mouse Visual Cortex Using a Thinned-skull Preparation

Authors: Emily A. Kelly, Ania K. Majewska.

Institutions: University of Rochester.

In vivo imaging using two-photon laser scanning microscopy (2PLSM) allows the study of living cells and neuronal processes in the intact brain. The technique presented here allows the imaging of the same area of the brain at several time points (chronic imaging) with microscopic resolution allowing the tracking of dendritic spines which are the small structures that represent the majority of postsynaptic excitatory sites in the CNS. The ability to clearly resolve fine cortical structures over several time points has many advantages, specifically in the study of brain plasticity in which morphological changes at synapses and circuit remodeling may help explain underlying mechanisms. In this video and supplementary material, we show a protocol for chronic in vivo imaging of the intact brain using a thinned-skull preparation. The thinned-skull preparation is a minimally invasive approach, which avoids potential damage to the dura and/or cortex, thus reducing the onset of an inflammatory response. When this protocol is performed correctly, it is possible to clearly monitor changes in dendritic spine characteristics in the intact brain over a prolonged period of time.

Neuronal morphology plays a significant role in determining how neurons function and communicate1-3. Specifically, it affects the ability of neurons to receive inputs from other cells2 and contributes to the propagation of action potentials4,5. The morphology of the neurites also affects how information is processed. The diversity of dendrite morphologies facilitate local and long range signaling and allow individual neurons or groups of neurons to carry out specialized functions within the neuronal network6,7. Alterations in dendrite morphology, including fragmentation of dendrites and changes in branching patterns, have been observed in a number of disease states, including Alzheimer's disease8, schizophrenia9,10, and mental retardation11. The ability to both understand the factors that shape dendrite morphologies and to identify changes in dendrite morphologies is essential in the understanding of nervous system function and dysfunction.
Neurite morphology is often analyzed by Sholl analysis and by counting the number of neurites and the number of branch tips. This analysis is generally applied to dendrites, but it can also be applied to axons. Performing this analysis by hand is both time consuming and inevitably introduces variability due to experimenter bias and inconsistency. The Bonfire program is a semi-automated approach to the analysis of dendrite and axon morphology that builds upon available open-source morphological analysis tools. Our program enables the detection of local changes in dendrite and axon branching behaviors by performing Sholl analysis on subregions of the neuritic arbor. For example, Sholl analysis is performed on both the neuron as a whole as well as on each subset of processes (primary, secondary, terminal, root, etc.) Dendrite and axon patterning is influenced by a number of intracellular and extracellular factors, many acting locally. Thus, the resulting arbor morphology is a result of specific processes acting on specific neurites, making it necessary to perform morphological analysis on a smaller scale in order to observe these local variations12.
The Bonfire program requires the use of two open-source analysis tools, the NeuronJ plugin to ImageJ and NeuronStudio. Neurons are traced in ImageJ, and NeuronStudio is used to define the connectivity between neurites. Bonfire contains a number of custom scripts written in MATLAB (MathWorks) that are used to convert the data into the appropriate format for further analysis, check for user errors, and ultimately perform Sholl analysis. Finally, data are exported into Excel for statistical analysis. A flow chart of the Bonfire program is shown in Figure 1.

Institutions: Medical College of Wisconsin, Stanford University School of Medicine, Medical College of Wisconsin, Hong Kong University, Johns Hopkins University School of Medicine, Medical College of Wisconsin.

There is an urgent need to develop approaches for repairing the damaged heart, discovering new therapeutic drugs that do not have toxic effects on the heart, and improving strategies to accurately model heart disease. The potential of exploiting human induced pluripotent stem cell (hiPSC) technology to generate cardiac muscle “in a dish” for these applications continues to generate high enthusiasm. In recent years, the ability to efficiently generate cardiomyogenic cells from human pluripotent stem cells (hPSCs) has greatly improved, offering us new opportunities to model very early stages of human cardiac development not otherwise accessible. In contrast to many previous methods, the cardiomyocyte differentiation protocol described here does not require cell aggregation or the addition of Activin A or BMP4 and robustly generates cultures of cells that are highly positive for cardiac troponin I and T (TNNI3, TNNT2), iroquois-class homeodomain protein IRX-4 (IRX4), myosin regulatory light chain 2, ventricular/cardiac muscle isoform (MLC2v) and myosin regulatory light chain 2, atrial isoform (MLC2a) by day 10 across all human embryonic stem cell (hESC) and hiPSC lines tested to date. Cells can be passaged and maintained for more than 90 days in culture. The strategy is technically simple to implement and cost-effective. Characterization of cardiomyocytes derived from pluripotent cells often includes the analysis of reference markers, both at the mRNA and protein level. For protein analysis, flow cytometry is a powerful analytical tool for assessing quality of cells in culture and determining subpopulation homogeneity. However, technical variation in sample preparation can significantly affect quality of flow cytometry data. Thus, standardization of staining protocols should facilitate comparisons among various differentiation strategies. Accordingly, optimized staining protocols for the analysis of IRX4, MLC2v, MLC2a, TNNI3, and TNNT2 by flow cytometry are described.

Dopaminergic neurons represent less than 1% of the total number of neurons in the brain. This low amount of neurons regulates important brain functions such as motor control, motivation, and working memory. Nigrostriatal dopaminergic neurons selectively degenerate in Parkinson's disease (PD). This progressive neuronal loss is unequivocally associated with the motors symptoms of the pathology (bradykinesia, resting tremor, and muscular rigidity). The main agent responsible of dopaminergic neuron degeneration is still unknown. However, these neurons appear to be extremely vulnerable in diverse conditions. Primary cultures constitute one of the most relevant models to investigate properties and characteristics of dopaminergic neurons. These cultures can be submitted to various stress agents that mimic PD pathology and to neuroprotective compounds in order to stop or slow down neuronal degeneration. The numerous transgenic mouse models of PD that have been generated during the last decade further increased the interest of researchers for dopaminergic neuron cultures. Here, the video protocol focuses on the delicate dissection of embryonic mouse brains. Precise excision of ventral mesencephalon is crucial to obtain neuronal cultures sufficiently rich in dopaminergic cells to allow subsequent studies. This protocol can be realized with embryonic transgenic mice and is suitable for immunofluorescence staining, quantitative PCR, second messenger quantification, or neuronal death/survival assessment.

Institutions: University of Michigan, University of Michigan, University of Michigan, University of Michigan, University of Michigan.

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of the Drosophila larva makes it an attractive model organism for live imaging studies. However, an important challenge for live imaging techniques is to noninvasively immobilize and position an animal on the microscope. This protocol presents a simple and easy to use method for immobilizing and imaging Drosophila larvae on a polydimethylsiloxane (PDMS) microfluidic device, which we call the 'larva chip'. The larva chip is comprised of a snug-fitting PDMS microchamber that is attached to a thin glass coverslip, which, upon application of a vacuum via a syringe, immobilizes the animal and brings ventral structures such as the nerve cord, segmental nerves, and body wall muscles, within close proximity to the coverslip. This allows for high-resolution imaging, and importantly, avoids the use of anesthetics and chemicals, which facilitates the study of a broad range of physiological processes. Since larvae recover easily from the immobilization, they can be readily subjected to multiple imaging sessions. This allows for longitudinal studies over time courses ranging from hours to days. This protocol describes step-by-step how to prepare the chip and how to utilize the chip for live imaging of neuronal events in 3rd instar larvae. These events include the rapid transport of organelles in axons, calcium responses to injury, and time-lapse studies of the trafficking of photo-convertible proteins over long distances and time scales. Another application of the chip is to study regenerative and degenerative responses to axonal injury, so the second part of this protocol describes a new and simple procedure for injuring axons within peripheral nerves by a segmental nerve crush.

In both mammals and insects neuronal information is processed in different higher and lower order brain centers. These centers are coupled via convergent and divergent anatomical connections including feed forward and feedback wiring. Furthermore, information of the same origin is partially sent via parallel pathways to different and sometimes into the same brain areas. To understand the evolutionary benefits as well as the computational advantages of these wiring strategies and especially their temporal dependencies on each other, it is necessary to have simultaneous access to single neurons of different tracts or neuropiles in the same preparation at high temporal resolution. Here we concentrate on honeybees by demonstrating a unique extracellular long term access to record multi unit activity at two subsequent neuropiles1, the antennal lobe (AL), the first olfactory processing stage and the mushroom body (MB), a higher order integration center involved in learning and memory formation, or two parallel neuronal tracts2 connecting the AL with the MB. The latter was chosen as an example and will be described in full. In the supporting video the construction and permanent insertion of flexible multi channel wire electrodes is demonstrated. Pairwise differential amplification of the micro wire electrode channels drastically reduces the noise and verifies that the source of the signal is closely related to the position of the electrode tip. The mechanical flexibility of the used wire electrodes allows stable invasive long term recordings over many hours up to days, which is a clear advantage compared to conventional extra and intracellular in vivo recording techniques.

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Authors: Mean-Hwan Kim, Evan Vickers, Henrique von Gersdorff.

Institutions: Oregon Health and Science University.

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate retina. Two classes of retinal neurons, photoreceptors and bipolar cells, accomplish this by using ribbon-type active zones, which enable sustained and high-throughput neurotransmitter release over long time periods. ON-type mixed bipolar cell (Mb) terminals in the goldfish retina, which depolarize to light stimuli and receive mixed rod and cone photoreceptor input, are suitable for the study of ribbon-type synapses both due to their large size (~10-12 μm diameter) and to their numerous lateral and reciprocal synaptic connections with amacrine cell dendrites. Direct access to Mb bipolar cell terminals in goldfish retinal slices with the patch-clamp technique allows the measurement of presynaptic Ca2+ currents, membrane capacitance changes, and reciprocal synaptic feedback inhibition mediated by GABAA and GABAC receptors expressed on the terminals. Presynaptic membrane capacitance measurements of exocytosis allow one to study the short-term plasticity of excitatory neurotransmitter release 14,15. In addition, short-term and long-term plasticity of inhibitory neurotransmitter release from amacrine cells can also be investigated by recordings of reciprocal feedback inhibition arriving at the Mb terminal 21. Over short periods of time (e.g. ~10 s), GABAergic reciprocal feedback inhibition from amacrine cells undergoes paired-pulse depression via GABA vesicle pool depletion 11. The synaptic dynamics of retinal microcircuits in the inner plexiform layer of the retina can thus be directly studied.
The brain-slice technique was introduced more than 40 years ago but is still very useful for the investigation of the electrical properties of neurons, both at the single cell soma, single dendrite or axon, and microcircuit synaptic level 19. Tissues that are too small to be glued directly onto the slicing chamber are often first embedded in agar (or placed onto a filter paper) and then sliced 20, 23, 18, 9. In this video, we employ the pre-embedding agar technique using goldfish retina. Some of the giant bipolar cell terminals in our slices of goldfish retina are axotomized (axon-cut) during the slicing procedure. This allows us to isolate single presynaptic nerve terminal inputs, because recording from axotomized terminals excludes the signals from the soma-dendritic compartment. Alternatively, one can also record from intact Mb bipolar cells, by recording from terminals attached to axons that have not been cut during the slicing procedure. Overall, use of this experimental protocol will aid in studies of retinal synaptic physiology, microcircuit functional analysis, and synaptic transmission at ribbon synapses.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

Visualizing the Effects of a Positive Early Experience, Tactile Stimulation, on Dendritic Morphology and Synaptic Connectivity with Golgi-Cox Staining

Authors: Richelle Mychasiuk, Robbin Gibb, Bryan Kolb.

Institutions: University of Lethbridge.

To generate longer-term changes in behavior, experiences must be producing stable changes in neuronal morphology and synaptic connectivity. Tactile stimulation is a positive early experience that mimics maternal licking and grooming in the rat. Exposing rat pups to this positive experience can be completed easily and cost-effectively by using highly accessible materials such as a household duster. Using a cross-litter design, pups are either stroked or left undisturbed, for 15 min, three times per day throughout the perinatal period. To measure the neuroplastic changes related to this positive early experience, Golgi-Cox staining of brain tissue is utilized. Owing to the fact that Golgi-Cox impregnation stains a discrete number of neurons rather than all of the cells, staining of the rodent brain with Golgi-Cox solution permits the visualization of entire neuronal elements, including the cell body, dendrites, axons, and dendritic spines. The staining procedure is carried out over several days and requires that the researcher pay close attention to detail. However, once staining is completed, the entire brain has been impregnated and can be preserved indefinitely for ongoing analysis. Therefore, Golgi-Cox staining is a valuable resource for studying experience-dependent plasticity.

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. The optic nerve can be accessed within the orbit of the eye and completely transected (axotomized), cutting the axons of the entire RGC population. Optic nerve transection is a reproducible model of apoptotic neuronal cell death in the adult CNS 1-4. This model is particularly attractive because the vitreous chamber of the eye acts as a capsule for drug delivery to the retina, permitting experimental manipulations via intraocular injections. The diffusion of chemicals through the vitreous fluid ensures that they act upon the entire RGC population. Viral vectors, plasmids or short interfering RNAs (siRNAs) can also be delivered to the vitreous chamber in order to infect or transfect retinal cells 5-12. The high tropism of Adeno-Associated Virus (AAV) vectors is beneficial to target RGCs, with an infection rate approaching 90% of cells near the injection site 6, 7, 13-15. Moreover, RGCs can be selectively transfected by applying siRNAs, plasmids, or viral vectors to the cut end of the optic nerve 16-19 or injecting vectors into their target the superior colliculus 10. This allows researchers to study apoptotic mechanisms in the injured neuronal population without confounding effects on other bystander neurons or surrounding glia. RGC apoptosis has a characteristic time-course whereby cell death is delayed 3-4 days postaxotomy, after which the cells rapidly degenerate. This provides a window for experimental manipulations directed against pathways involved in apoptosis. Manipulations that directly target RGCs from the transected optic nerve stump are performed at the time of axotomy, immediately after cutting the nerve. In contrast, when substances are delivered via an intraocular route, they can be injected prior to surgery or within the first 3 days after surgery, preceding the initiation of apoptosis in axotomized RGCs. In the present article, we demonstrate several methods for experimental manipulations after optic nerve transection.

Phagocytosis is a process in which a cell engulfs material (entire cell, parts of a cell, debris, etc.) in its surrounding extracellular environment and subsequently digests this material, commonly through lysosomal degradation. Microglia are the resident immune cells of the central nervous system (CNS) whose phagocytic function has been described in a broad range of conditions from neurodegenerative disease (e.g., beta-amyloid clearance in Alzheimer’s disease) to development of the healthy brain (e.g., synaptic pruning)1-6. The following protocol is an engulfment assay developed to visualize and quantify microglia-mediated engulfment of presynaptic inputs in the developing mouse retinogeniculate system7. While this assay was used to assess microglia function in this particular context, a similar approach may be used to assess other phagocytes throughout the brain (e.g., astrocytes) and the rest of the body (e.g., peripheral macrophages) as well as other contexts in which synaptic remodeling occurs (e.g. ,brain injury/disease).

Neuronal axons use specific mechanisms to mediate extension, maintain integrity, and induce degeneration. An appropriate balance of these events is required to shape functional neuronal circuits. The protocol described here explains how to use cell culture inserts bearing a porous membrane (filter) to obtain large amounts of pure axonal preparations suitable for examination by conventional biochemical or immunocytochemical techniques. The functionality of these filter inserts will be demonstrated with models of developmental pruning and Wallerian degeneration, using explants of embryonic dorsal root ganglion. Axonal integrity and function is compromised in a wide variety of neurodegenerative pathologies. Indeed, it is now clear that axonal dysfunction appears much earlier in the course of the disease than neuronal soma loss in several neurodegenerative diseases, indicating that axonal-specific processes are primarily targeted in these disorders. By obtaining pure axonal samples for analysis by molecular and biochemical techniques, this technique has the potential to shed new light into mechanisms regulating the physiology and pathophysiology of axons. This in turn will have an impact in our understanding of the processes that drive degenerative diseases of the nervous system.

GABAergic inhibitory interneurons play a central role within neuronal circuits of the brain. Interneurons comprise a small subset of the neuronal population (10-20%), but show a high level of physiological, morphological, and neurochemical heterogeneity, reflecting their diverse functions. Therefore, investigation of interneurons provides important insights into the organization principles and function of neuronal circuits. This, however, requires an integrated physiological and neuroanatomical approach for the selection and identification of individual interneuron types. Whole-cell patch-clamp recording from acute brain slices of transgenic animals, expressing fluorescent proteins under the promoters of interneuron-specific markers, provides an efficient method to target and electrophysiologically characterize intrinsic and synaptic properties of specific interneuron types. Combined with intracellular dye labeling, this approach can be extended with post-hoc morphological and immunocytochemical analysis, enabling systematic identification of recorded neurons. These methods can be tailored to suit a broad range of scientific questions regarding functional properties of diverse types of cortical neurons.

Institutions: Northwestern University Feinberg School of Medicine, Northwestern University Feinberg School of Medicine.

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic
compartment of synapses. These structures are rich in actin and have been shown to be highly dynamic. In response to classical Hebbian plasticity
as well as neuromodulatory signals, dendritic spines can change shape and number, which is thought to be critical for the refinement of neural
circuits and the processing and storage of information within the brain. Within dendritic spines, a complex network of proteins link extracellular
signals with the actin cyctoskeleton allowing for control of dendritic spine morphology and number. Neuropathological studies have demonstrated that
a number of disease states, ranging from schizophrenia to autism spectrum disorders, display abnormal dendritic spine morphology or numbers.
Moreover, recent genetic studies have identified mutations in numerous genes that encode synaptic proteins, leading to suggestions that these
proteins may contribute to aberrant spine plasticity that, in part, underlie the pathophysiology of these disorders. In order to study the potential
role of these proteins in controlling dendritic spine morphologies/number, the use of cultured cortical neurons offers several advantages. Firstly,
this system allows for high-resolution imaging of dendritic spines in fixed cells as well as time-lapse imaging of live cells. Secondly, this in
vitro system allows for easy manipulation of protein function by expression of mutant proteins, knockdown by shRNA constructs, or pharmacological
treatments. These techniques allow researchers to begin to dissect the role of disease-associated proteins and to predict how mutations of these
proteins may function in vivo.

Inhibitory neurons act in the central nervous system to regulate the dynamics and spatio-temporal co-ordination of neuronal networks. GABA (γ-aminobutyric acid) is the predominant inhibitory neurotransmitter in the brain. It is released from the presynaptic terminals of inhibitory neurons within highly specialized intercellular junctions known as synapses, where it binds to GABAA receptors (GABAARs) present at the plasma membrane of the synapse-receiving, postsynaptic neurons. Activation of these GABA-gated ion channels leads to influx of chloride resulting in postsynaptic potential changes that decrease the probability that these neurons will generate action potentials.
During development, diverse types of inhibitory neurons with distinct morphological, electrophysiological and neurochemical characteristics have the ability to recognize their target neurons and form synapses which incorporate specific GABAARs subtypes. This principle of selective innervation of neuronal targets raises the question as to how the appropriate synaptic partners identify each other.
To elucidate the underlying molecular mechanisms, a novel in vitro co-culture model system was established, in which medium spiny GABAergic neurons, a highly homogenous population of neurons isolated from the embryonic striatum, were cultured with stably transfected HEK293 cell lines that express different GABAAR subtypes. Synapses form rapidly, efficiently and selectively in this system, and are easily accessible for quantification. Our results indicate that various GABAAR subtypes differ in their ability to promote synapse formation, suggesting that this reduced in vitro model system can be used to reproduce, at least in part, the in vivo conditions required for the recognition of the appropriate synaptic partners and formation of specific synapses. Here the protocols for culturing the medium spiny neurons and generating HEK293 cells lines expressing GABAARs are first described, followed by detailed instructions on how to combine these two cell types in co-culture and analyze the formation of synaptic contacts.

Multi-photon fluorescence microscopy has enabled the analysis of morphological and physiological parameters of brain cells in the intact tissue with high spatial and temporal resolution. Combined with electrophysiology, it is widely used to study activity-related calcium signals in small subcellular compartments such as dendrites and dendritic spines. In addition to calcium transients, synaptic activity also induces postsynaptic sodium signals, the properties of which are only marginally understood. Here, we describe a method for combined whole-cell patch-clamp and multi-photon sodium imaging in cellular micro domains of central neurons. Furthermore, we introduce a modified procedure for ultra-violet (UV)-light-induced uncaging of glutamate, which allows reliable and focal activation of glutamate receptors in the tissue. To this end, whole-cell recordings were performed on Cornu Ammonis subdivision 1 (CA1) pyramidal neurons in acute tissue slices of the mouse hippocampus. Neurons were filled with the sodium-sensitive fluorescent dye SBFI through the patch-pipette, and multi-photon excitation of SBFI enabled the visualization of dendrites and adjacent spines. To establish UV-induced focal uncaging, several parameters including light intensity, volume affected by the UV uncaging beam, positioning of the beam as well as concentration of the caged compound were tested and optimized. Our results show that local perfusion with caged glutamate (MNI-Glutamate) and its focal UV-uncaging result in inward currents and sodium transients in dendrites and spines. Time course and amplitude of both inward currents and sodium signals correlate with the duration of the uncaging pulse. Furthermore, our results show that intracellular sodium signals are blocked in the presence of blockers for ionotropic glutamate receptors, demonstrating that they are mediated by sodium influx though this pathway. In summary, our method provides a reliable tool for the investigation of intracellular sodium signals induced by focal receptor activation in intact brain tissue.

Institutions: University of Helsinki, Neurotar LTD, University of Eastern Finland, University of Helsinki.

It is widely acknowledged that the use of general anesthetics can undermine the relevance of electrophysiological or microscopical data obtained from a living animal’s brain. Moreover, the lengthy recovery from anesthesia limits the frequency of repeated recording/imaging episodes in longitudinal studies. Hence, new methods that would allow stable recordings from non-anesthetized behaving mice are expected to advance the fields of cellular and cognitive neurosciences. Existing solutions range from mere physical restraint to more sophisticated approaches, such as linear and spherical treadmills used in combination with computer-generated virtual reality. Here, a novel method is described where a head-fixed mouse can move around an air-lifted mobile homecage and explore its environment under stress-free conditions. This method allows researchers to perform behavioral tests (e.g., learning, habituation or novel object recognition) simultaneously with two-photon microscopic imaging and/or patch-clamp recordings, all combined in a single experiment. This video-article describes the use of the awake animal head fixation device (mobile homecage), demonstrates the procedures of animal habituation, and exemplifies a number of possible applications of the method.

Institutions: University of Wuerzburg, Max Planck Institute of Neurobiology, Martinsried, Ludwig-Maximilians University of Munich.

Visualization of calcium dynamics is important to understand the role of calcium in cell physiology. To examine calcium dynamics, synthetic fluorescent Ca2+ indictors have become popular. Here we demonstrate TED (= targeted-esterase induced dye loading), a method to improve the release of Ca2+ indicator dyes in the ER lumen of different cell types. To date, TED was used in cell lines, glial cells, and neurons in vitro. TED bases on efficient, recombinant targeting of a high carboxylesterase activity to the ER lumen using vector-constructs that express Carboxylesterases (CES). The latest TED vectors contain a core element of CES2 fused to a red fluorescent protein, thus enabling simultaneous two-color imaging. The dynamics of free calcium in the ER are imaged in one color, while the corresponding ER structure appears in red. At the beginning of the procedure, cells are transduced with a lentivirus. Subsequently, the infected cells are seeded on coverslips to finally enable live cell imaging. Then, living cells are incubated with the acetoxymethyl ester (AM-ester) form of low-affinity Ca2+ indicators, for instance Fluo5N-AM, Mag-Fluo4-AM, or Mag-Fura2-AM. The esterase activity in the ER cleaves off hydrophobic side chains from the AM form of the Ca2+ indicator and a hydrophilic fluorescent dye/Ca2+ complex is formed and trapped in the ER lumen. After dye loading, the cells are analyzed at an inverted confocal laser scanning microscope. Cells are continuously perfused with Ringer-like solutions and the ER calcium dynamics are directly visualized by time-lapse imaging. Calcium release from the ER is identified by a decrease in fluorescence intensity in regions of interest, whereas the refilling of the ER calcium store produces an increase in fluorescence intensity. Finally, the change in fluorescent intensity over time is determined by calculation of ΔF/F0.

Cortical development involves complex interactions between neurons and non-neuronal elements including precursor cells, blood vessels, meninges and associated extracellular matrix. Because they provide a suitable organotypic environment, cortical slice explants are often used to investigate those interactions that control neuronal differentiation and development. Although beneficial, the slice explant model can suffer from drawbacks including aberrant cellular lamination and migration. Here we report a whole cerebral hemisphere explant system for studies of early cortical development that is easier to prepare than cortical slices and shows consistent organotypic migration and lamination. In this model system, early lamination and migration patterns proceed normally for a period of two days in vitro, including the period of preplate splitting, during which prospective cortical layer six forms. We then developed an ex utero electroporation (EUEP) approach that achieves ~80% success in targeting GFP expression to neurons developing in the dorsal medial cortex.
The whole hemisphere explant model makes early cortical development accessible for electroporation, pharmacological intervention and live imaging approaches. This method avoids the survival surgery required of in utero electroporation (IUEP) approaches while improving both transfection and areal targeting consistency. This method will facilitate experimental studies of neuronal proliferation, migration and differentiation.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

Authors: Onkar S. Dhande, Michael C. Crair.

Institutions: Yale University, Baylor College of Medicine.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6.
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

We present a protocol for in vivo imaging of cortical tissue using a deep-brain imaging probe in the shape of a microprism. Microprisms are 1-mm in size and have a reflective coating on the hypotenuse to allow internal reflection of excitation and emission light. The microprism probe simultaneously images multiple cortical layers with a perspective typically seen only in slice preparations. Images are collected with a large field-of-view (~900 μm). In addition, we provide details on the non-survival surgical procedure and microscope setup. Representative results include images of layer V pyramidal neurons from Thy-1 YFP-H mice showing their apical dendrites extending through the superficial cortical layer and extending into tufts. Resolution was sufficient to image dendritic spines near the soma of layer V neurons. A tail-vein injection of fluorescent dye reveals the intricate network of blood vessels in the cortex. Line-scanning of red blood cells (RBCs) flowing through the capillaries reveals RBC velocity and flux rates can be obtained. This novel microprism probe is an elegant, yet powerful new method of visualizing deep cellular structures and cortical function in vivo.

Institutions: University of California, Los Angeles, University of California, Los Angeles.

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.
Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Traditionally the vertical slice and the whole-mount preparation of the retina have been used to study the function of retinal circuits. However, many of retinal neurons, such as amacrine cells, expand their dendrites horizontally, so that the morphology of the cells is supposed to be severely damaged in the vertical slices. In the whole-mount preparation, especially for patch-clamp recordings, retinal neurons in the middle layer are not easily accessible due to the extensive coverage of glial cell (Mueller cell) s endfeets. Here, we describe the novel slicing method to preserve the dendritic morphology of retinal neurons intact. The slice was made horizontally at the inner layer of the retina using a vibratome slicer after the retina was embedded in the low-temperature melting agarose gel. In this horizontal slice preparation of the retina, we studied the function of retinal neurons compared with their morphology, by using patch-clamp recording, calcium imaging technique, immunocytochemistry, and single-cell RT-PCR.

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